DNA Replication in Archaea, the Third Domain of Lifecdn.intechopen.com/pdfs/.../InTech-Dna_replication_in_archaea_the_third... · logues of many eukaryotic replication proteins,

Chapter 4

DNA Replication in Archaea, the Third Domain of Life

Yoshizumi Ishino and Sonoko Ishino

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53986

1. Introduction

The accurate duplication and transmission of genetic information are essential and cruciallyimportant for living organisms. The molecular mechanism of DNA replication has been oneof the central themes of molecular biology, and continuous efforts to elucidate the precisemolecular mechanism of DNA replication have been made since the discovery of the doublehelix DNA structure in 1953 [1]. The protein factors that function in the DNA replicationprocess, have been identified to date in the three domains of life (Figure 1).

Figure 1. Stage of DNA replication

© 2013 Ishino and Ishino; licensee InTech. This is an open access article distributed under the terms of theCreative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permitsunrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Archaea Eukaryota Bacteria

initiation

origin recognition Cdc6/Orc1 ORC DnaA

DNA unwinding Cdc6/Orc1

MCM

GINS

Cdc6

Cdt1

MCM

GINS

Cdc45

DnaC

DnaB

primer synthesis DNA primase Pol α / primase DnaG

elongation

DNA synthesis family B

DNA polymerase

(Pol B)

family D

DNA polymerase

(Pol D)

family B

DNA polymerase

(Pol δ) (Pol ε)

family C

DNA polymerase

(Pol III)

clamp loader

(RFC)

clamp

(PCNA)

clamp loader

(RFC)

clamp

(PCNA)

clamp loader

(γ-complex)

clamp

(β-clamp)

maturation Fen1

Dna2

DNA ligase

FEN1

DNA2

DNA ligase

Pol I

RNaseH

DNA ligase

Table 1. The proteins involved in DNA replication from the three domains of life

Archaea, the third domain of life, is a very interesting living organism to study from the as‐pects of molecular and evolutional biology. Rapid progress of whole genome sequence anal‐yses has allowed us to perform comparative genomic studies. In addition, recent microbialecology has revealed that archaeal organisms inhabit not only extreme environments, but al‐so more ordinary habitats. In these situations, archaeal biology is among the most exciting ofresearch fields. Archaeal cells have a unicellular ultrastructure without a nucleus, resem‐bling bacterial cells, but the proteins involved in the genetic information processing path‐ways, including DNA replication, transcription, and translation, share strong similaritieswith those of eukaryotes. Therefore, most of the archaeal proteins were identified as homo‐logues of many eukaryotic replication proteins, including ORC (origin recognition complex),Cdc6, GINS (Sld5-Psf1-Psf2-Psf3), MCM (minichromosome maintenance), RPA (replicationprotein A), PCNA (proliferating cell nuclear antigen), RFC (replication factor C), FEN1 (flapendonuclease 1), in addition to the eukaryotic primase, DNA polymerase, and DNA ligase;these are obviously different from bacterial proteins (Table 1) and these proteins were bio‐chemically characterized [2-4]. Their similarities indicate that the DNA replication machi‐

The Mechanisms of DNA Replication92

neries of Archaea and Eukaryota evolved from a common ancestor, which was differentfrom that of Bacteria [5]. Therefore, the archaeal organisms are good models to elucidate thefunctions of each component of the eukaryotic type replication machinery complex. Genom‐ic and comparative genomic research with archaea is made easier by the fact that the ge‐nome size and the number of genes of archaea are much smaller than those of eukaryotes.The archaeal replication machinery is probably a simplified form of that in eukaryotes. Onthe other hand, it is also interesting that the circular genome structure is conserved in Bacte‐ria and Archaea and is different from the linear form of eukaryotic genomes. These featureshave encouraged us to study archaeal DNA replication, in the hopes of gaining fundamentalinsights into this molecular mechanism and its machinery from an evolutional perspective.The study of bacterial DNA replication at a molecular level started in about 1960, and theneukaryotic studies followed since 1980. Because Archaea was recognized as the third do‐main of life later, the archaeal DNA replication research became active after 1990. With in‐creasing the available total genome sequences, the progress of research on archaeal DNAreplication has been rapid, and the depth of our knowledge of archaeal DNA replication hasalmost caught up with those of the bacterial and eukaryotic research fields. In this chapter,we will summarize the current knowledge of DNA replication in Archaea.

2. Replication origin

The basic mechanism of DNA replication was predicted as “replicon theory” by Jacob et al.[6]. They proposed that an initiation factor recognizes the replicator, now referred to as areplication origin, to start replication of the chromosomal DNA. Then, the replication originof E. coli DNA was identified as oriC (origin of chromosome). The archaeal replication originwas identified in the Pyrococcus abyssi in 2001 as the first archaeal replication origin. The ori‐gin was located just upstream of the gene encoding the Cdc6 and Orc1-like sequences in thePyrococcus genome [7]. We discovered a gene encoding an amino acid sequence that boresimilarity to those of both eukaryotic Cdc6 and Orc1, which are the eukaryotic initiators. Af‐ter confirming that this protein actually binds to the oriC region on the chromosomal DNAwe named the gene product Cdc6/Orc1 due to its roughly equal homology with regions ofeukaryotic Orc1 and Cdc6, [7]. The gene consists of an operon with the gene encoding DNApolymerase D (it was originally called Pol II, as the second DNA polymerase from Pyrococ‐cus furiosus) in the genome [8]. A characteristic of the oriC is the conserved 13 bp repeats, aspredicted earlier by bioinformatics [9], and two of the repeats are longer and surround apredicted DUE (DNA unwinding element) with an AT-rich sequence in Pyrococcus genomes(Figure 2) [10]. The longer repeated sequence was designated as an ORB (Origin RecognitionBox), and it was actually recognized by Cdc6/Orc1 in a Sulfolobus solfataricus study [11]. The13 base repeat is called a miniORB, as a minimal version of ORB. A whole genome microar‐ray analysis of P. abyssi showed that the Cdc6/Orc1 binds to the oriC region with extremespecificity, and the specific binding of the highly purified P. furiosus Cdc6/Orc1 to ORB andminiORB was confirmed in vitro [12]. It has to be noted that multiple origins were identifiedin the Sulfolobus genomes. It is now well recognized that Sulfolobus has three origins and

DNA Replication in Archaea, the Third Domain of Lifehttp://dx.doi.org/10.5772/53986

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they work at the same time in the cell cycle [11, 13-16]. Analysis of the mechanism of howthe multiple origins are utilized for genome replication is an interesting subject in the re‐search field of archaeal DNA replication. The main questions are how the initiation of repli‐cation from multiple origins is regulated and how the replication forks progress after thecollision of two forks from opposite directions.

Figure 2. The oriC region in Pyrococcus genome. The region surrounding oriC is presented schematically. The ORB1and ORB2 are indicated by large arrow, and the mini-ORB repeats are indicated by small arrowheads. DUE is indicatedin red. The unwinding site, determined by in vitro analysis, is indicated in orange. The transition site is indicated bygreen arrows. The cdc6/orc1 gene located in downstream is drawn by gray arrow.

3. How does Cdc6/Orc1 recognize oriC?

An important step in characterizing the initiation of DNA replication in Archaea is to under‐stand how the Cdc6/Orc1 protein recognizes the oriC region. Based upon amino acid se‐quence alignments, the archaeal Cdc6/Orc1 proteins belong to the AAA+ family of proteins.The crystal structures of the Cdc6/Orc1 protein from Pyrobaculum aerophilum [17] and one ofthe two Cdc6/Orc1 proteins, ORC2 from Aeropyrum pernix (the two homologs in this organ‐ism are called ORC1 and ORC2 by the authors) [18] were determined. These Cdc6/Orc1 pro‐teins consist of three structural domains. Domains I and II adopt a fold found in the AAA+

family proteins. A winged helix (WH) fold, which is present in a number of DNA bindingproteins, is found in the domain III. There are four ORBs arranged in pairs on both sides ofthe DUE in the oriC region of A. pernix, and ORC1 binds to each ORB as a dimer. A mecha‐nism was proposed in which ORC1 binds to all four ORBs to introduce a higher-order as‐sembly for unwinding of the DUE with alterations in both topology and superhelicity [19].Furthermore, the crystal structures of S. solfataricus Cdc6-1 and Cdc6-3 (two of the three


Cdc6/Orc1 proteins in this organism) forming a heterodimer bound to ori2 DNA (one of thethree origins in this organism) [20], and that of A. pernix ORC1 bound to an origin sequence[21] were determined. These studies revealed that both the N-terminal AAA+ ATPase do‐main (domain I+II) and C-terminal WH domain (domain III) contribute to origin DNA bind‐ing, and the structural information not only defined the polarity of initiator assembly on theorigin but also indicated the induction of substantial distortion, which probably triggers theunwinding of the duplex DNA to start replication, into the DNA strands. These structuraldata also provided the detailed interaction mode between the initiator protein and the oriCDNA. Mutational analyses of the Methanothermobactor thermautotrophicus Cdc6-1 protein re‐vealed the essential interaction between an arginine residue conserved in the archaeal Cdc6/Orc1 and an invariant guanine in the ORB sequence [22].

P. furiosus Cdc6/Orc1 is difficult to purify in a soluble form. A specific site in the oriC to startunwinding in vitro, was identified using the protein prepared by a denaturation-renatura‐tion procedure recently [23]. As shown in Figure 2, the local unwinding site is about 670 bpaway from the transition site between leading and lagging syntheses, which was determinedearlier by an in vivo replication initiation point (RIP) assay [10]. Although the details of thereplication machinery that must be established at the unwound site are not fully understoodin Archaea, it is expected to minimally include MCM, GINS, primase, PCNA, DNA poly‐merase, and RPA, as described below. The following P. furiosus studies revealed that the AT‐Pase activity of the Cdc6/Orc1 protein was completely suppressed by binding to DNAcontaining the ORB. Limited proteolysis and DNase I-footprint experiments suggested thatthe Cdc6/Orc1 protein changes its conformation on the ORB sequence in the presence ofATP. The physiological meaning of this conformational change has not been solved, but itshould have an important function to start the initiation process [24] as in the case of bacteri‐al DnaA protein. In addition, results from an in vitro recruiting assay indicated that MCM(Mcm protein complex), the replicative DNA helicase, is recruited onto the oriC region in aCdc6/Orc1-dependent, but not ATP-dependent, manner [24], as described below. However,this recruitment is not sufficient for the unwinding function of MCM, and some other func‐tion remains to be identified for the functional loading of this helicase to promote the pro‐gression of the DNA replication fork.

4. MCM helicase

After unwinding of the oriC region, the replicative helicase needs to remain loaded to pro‐vide continuous unwinding of double stranded DNA (dsDNA) as the replication forks prog‐ress bidirectionally. The MCM protein complex, consisting of six subunits (Mcm2, 3, 4, 5, 6,and 7), is known to be the replicative helicase “core” in eukaryotic cells [25]. The MCM fur‐ther interacts with Cdc45 and GINS, to form a ternary assembly referred to as the “CMGcomplex”, that is believed to be the functional helicase in eukaryotic cells (Figure 3) [26].However, this idea is still not universal for the eukaryotic replicative helicase.


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Figure 3. DNA-Unwinding complex in eukaryotes and archaea. The CMG complex is the replicative helicase for thetemplate DNA unwinding reaction in eukaryotes. The archaeal genomes contain the homologs of the Mcm and Ginsproteins, but a Cdc45 homolog has not been identified. Recent research suggests that a RecJ-like exonuclease GAN,which has weak sequence homology to that of Cdc45, may work as a helicase complex with MCM and GINS.

Most archaeal genomes appear to encode at least one Mcm homologue, and the helicase ac‐tivities of these proteins from several archaeal organisms have been confirmed in vitro[27-31]. In contrast to the eukaryotic MCM, the archaeal MCMs, consist of a homohexameror homo double hexamer, having distinct DNA helicase activity by themselves in vitro, andtherefore, these MCMs on their own may function as the replicative helicase in vivo. Thestructure-function relationships of the archaeal Mcms have been aggressively studied usingpurified proteins and site-directed mutagenesis [32]. An early report using the ChIP methodshowed that the P. abyssi Mcm protein preferentially binds to the origin in vivo in exponen‐tially growing cells [7, 12]. The P. furiosus MCM helicase does not display significant helicaseactivity in vitro. However, the DNA helicase activity was clearly stimulated by the additionof GINS (the Gins23-Gins51 complex), which is the homolog of the eukaryotic GINS com‐plex (described below in more detail). This result suggests that MCM works with other ac‐cessory factors to form a core complex in P. furiosus similar to the eukaryotic CMG complexas described above [31].

Some archaeal organisms have more than two Cdc6/Orc1 homologs. It was found that thetwo Cdc6/Orc1 homologs, Cdc6-1 and Cdc6-2, both inhibit the helicase activity of MCM inM. thermautotrophicus [33. 34]. Similarly, Cdc6-1 inhibits MCM activity in S. solfataricus [35].In contrast, the Cdc6-2 protein stimulates the helicase activity of MCM in Thermoplasma acid‐ophilum [36]. Functional interactions between Cdc6/Orc1 and Mcm proteins need to be inves‐tigated in greater detail to achieve a more comprehensive understanding of the conservationand diversity of the initiation mechanism in archaeal DNA replication.

Another interesting feature of DNA replication initiation is that several archaea have multi‐ple genes encoding Mcm homologs in their genomes. Based on the recent comprehensivegenomic analyses, thirteen archaeal species have more than one mcm gene. However, manyof the mcm genes in the archaeal genomes seem to reside within mobile elements, originat‐ing from viruses [37]. For example, two of the three genes in the Thermococcus kodakarensisgenome are located in regions where genetic elements have presumably been integrated[38]. The establishment of a genetic manipulation system for T. kodakarensis, is the first for ahyperthermophilic euryarchaeon [39. 40], and is advantageous for investigating the functionof these Mcm proteins. Two groups have recently performed gene disruption experimentsfor each mcm gene [41, 42]. These experiments revealed that the knock-out strains for mcm1


and mcm2 were easily isolated, but mcm3 could not be disrupted. Mcm3 is relatively abun‐dant in the T. kodakarensis cells. Furthermore, an in vitro experiment using purified Mcm pro‐teins showed that only Mcm3 forms a stable hexameric structure in solution. These resultssupport the contention that Mcm3 is the main helicase core protein in the normal DNA rep‐lication process in T. kodakarensis.

The functions of the other two Mcm proteins remain to be elucidated. The genes for Mcm1and Mcm2 are stably inherited, and their gene products may perform some important func‐tions in the DNA metabolism in T. kodakarensis. The DNA helicase activity of the recombi‐nant Mcm1 protein is strong in vitro, and a distinct amount of the Mcm1 protein is present inT. kodakarensis cells. Moreover, Mcm1 functionally interacts with the GINS complex from T.kodakarensis [42]. These observations strongly suggest that Mcm1 does participate in someaspect of DNA transactions, and may be substituted with Mcm3. Our immunoprecipitationexperiments showed that Mcm1 co-precipitated with Mcm3 and GINS, although they didnot form a heterohexameric complex [42], suggesting that Mcm1 is involved in the repli‐some or repairsome and shares some function in T. kodakarensis cells. Although western blotanalysis could not detect Mcm2 in the extract from exponentially growing T. kodakarensiscells [42], a RT-PCR experiment detected the transcript of the mcm2 gene in the cells (Ishinoet al., unpublished). The recombinant Mcm2 protein also has ATPase and helicase activitiesin vitro. [41] Therefore, the mcm2 gene is expressed under normal growth conditions andmay work in some process with a rapid turn over. Further experiments to measure the effi‐ciency of mcm2 gene transcription by quantitative PCR, as well as to assess the stability ofthe Mcm2 protein in the cell extract, are needed. Phenotypic analyses investigating the sensi‐tivities of the Δmcm1 and Δmcm2 mutant strains to DNA damage caused by various muta‐gens, as reported for other DNA repair-related genes in T. kodakarensis [43], may provide aclue to elucidate the functions of these Mcm proteins.

Methanococcus maripaludis S2 harbors four mcm genes in its genome, three of which seem tobe derived from phage, a shotgun proteomics study detected peptides originating fromthree out of the four mcm gene products [44]. Furthermore, the four gene products co-ex‐pressed in E. coli cells were co-purified in the same fraction [45]. These results suggest thatmultiple Mcm proteins are functional in the M. maripaludis cells.

5. Recruitment of Mcm to the oriC region

Another important question is how MCM is recruited onto the unwound region of oriC. Thedetailed loading mechanism of the MCM helicase has not been elucidated. It is believed thatarchaea utilize divergent mechanisms of MCM helicase assembly at the oriC [46].

An in vitro recruiting assay showed that P. furiosus MCM is recruited to the oriC DNA in aCdc6/Orc1-dependent manner [24]. This assay revealed that preloading Cdc6/Orc1 onto theORB DNA resulted in a clear reduction in MCM recruitment to the oriC region, suggestingthat free Cdc6/Orc1 is preferable as a helicase recruiter, to associate with MCM and bring itto oriC. It would be interesting to understand how the two tasks, origin recognition and


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MCM recruiting, are performed by the Cdc6/Orc1 protein, because the WH domain, whichprimarily recognizes and binds ORB, also has strong affinity for the Mcm protein. The as‐sembly of the Mcm protein onto the ORB DNA by the Walker A-motif mutant of P. furiosusCdc6/Orc1 occurred with the same efficiency as the wild type Cdc6/Orc1. The DNA bindingof P. furiosus Cdc6/Orc1 was not drastically different in the presence and absence of ATP, asin the case of the initiator proteins from Archaeoglobus fulgidus [28], S. solfataricus [11], and A.pernix [19]. Therefore, it is still not known whether the ATP binding and hydrolysis activityof Cdc6/Orc1 regulates the Mcm protein recruitment onto oriC in the cells.

One more important issue is the very low efficiency of the Mcm protein recruitment in thereported in vitro assay [24]. Quantification of the recruited Mcm protein by the in vitro assayshowed that less than one Mcm hexamer was recruited to the ORB. The linear DNA contain‐ing ORB1 and ORB2, used in the recruiting assay, may not be suitable to reconstitute thearchaeal DNA replication machinery and a template that more closely mimics the chromo‐somal DNA may be required. Additionally, it may be that as yet unidentified proteins arerequired to achieve efficient in vitro helicase loading in the P. furiosus cells. Finally, it willultimately be necessary to construct a more defined in vitro replication system to analyze theregulatory functions of Cdc6/Orc1 precisely during replication initiation.

In M. thermautotrophicus, the Cdc6-2 proteins can dissociate the Mcm multimers [47]. The ac‐tivity of Cdc6-2 might be required as the MCM helicase loader in this organism. The interac‐tion between Cdc6/Orc1 and Mcm is probably general. However, the effect of Cdc6/Orc1 onthe MCM helicase activity differs among various organisms, as described above. Some otherprotein factors may function in various archaea, for example a protein that is distantly relat‐ed to eukaryotic Cdt1, which plays a crucial role during MCM loading in Eukaryota, existsin some archaeal organisms, although its function has not been characterized yet [14].

6. GINS

The eukaryotic GINS complex was originally identified in Saccharomyces cerevisiae as essen‐tial protein factor for the initiation of DNA replication [48]. GINS consists of four differentproteins, Sld5, Psf1, Psf2, and Psf3 (therefore, GINS is an acronym for Japanese go-ichi-ni-san, meaning 5-1-2-3, after these four subunits). The amino acid sequences of the four subu‐nits in the GINS complex share some conservation, suggesting that they are ancestralparalogs [49]. However, most of the archaeal genomes have only one gene encoding thisfamily protein, and more interestingly, the Crenarchaeota and Euryarchaeota (the two majorsubdomains of Archaea) characteristically have two genes with sequences similar to Psf2and Psf3, and Sld5 and Psf1, respectively referred to as Gins23 and Gins51 [31, 49]. A Ginshomolog, designated as Gins23, was biochemically detected in S. solfataricus as the first Ginsprotein in Archaea, in a yeast two-hybrid screening for interaction partners of the Mcm pro‐tein, and another subunit, designated as Gins15, was identified by mass-spectrometry analy‐sis of an immunoaffinity-purified native GINS from an S. solfataricus cell extract. [50]. The S.solfataricus GINS, composed of two proteins, Gins23 and Gins15, forms a tetrameric struc‐


ture with a 2:2 molar ratio [50]. The GINS from P. furiosus, a complex of Gins23 and Gins51with a 2:2 ratio, was identified as the first euryarchaeal GINS [31]. Gins51 was preferredover Gins15 because of the order of the name of GINS.

The MCM2-7 hexamer was copurified in complex with Cdc45 and GINS from Drosophilamelanogaster embryo extracts and S. cerevisiae lysates, and the “CMG (Cdc45-MCM2-7-GINS)complex” (Figure 3), as described above, should be important for the function of the replica‐tive helicase. The CMG complex was also associated with the replication fork in Xenopus lae‐vis egg extracts, and a large molecular machine, containing Cdc45, GINS, and MCM2-7, wasproposed as the unwindosome to separate the DNA strands at the replication fork [51].Therefore, GINS must be a critical factor for not only the initiation process, but also the elon‐gation process in eukaryotic DNA replication. S. solfataricus GINS interacts with MCM andprimase, suggesting that GINS is involved in the replisome. The concrete function of GINSin the replisome remains to be determined. No stimulation or inhibition of either the heli‐case or primase activity was observed by the interaction with S. solfataricus GINS in vitro[50]. On the other hand, the DNA helicase activity of P. furiosus MCM is clearly stimulatedby the addition of the P. furiosus GINS complex, as described above [31].

In contrast to S. solfataricus and P. furiosus, which each express a Gins23 and Gins51, Thermo‐plasma acidophilum has a single Gins homolog, Gins51. The recombinant Gins51 protein fromT. acidophilum was confirmed to form a homotetramer by gel filtration and electron micro‐scopy analyses. Furthermore, a physical interaction between T. acidophilum Gins51 and Mcmwas detected by a surface plasmon resonance analysis (SPR). Although the T. acidophilumGins51 did not affect the helicase activity of its cognate MCM, when the equal ratio of eachmolecule was tested in vitro [52], an excess amount of Gins51 clearly stimulated the helicaseactivity (Ogino et al., unpublished). In the case of T. kodakarensis, the ATPase and helicaseactivities of MCM1 and MCM3 were clearly stimulated by T. kodakarensis GINS in vitro. It isinteresting that the helicase activity of MCM1 was stimulated more than that of MCM3.Physical interactions between the T. kodakarensis Gins and Mcm proteins were also detected[53]. These reports suggested that the MCM-GINS complex is a common part of the replica‐tive helicase in Archaea (Figure 3).

Recently, the crystal structure of the T. kodakarensis GINS tetramer, composed of Gins51 andGins23 was determined, and the structure was conserved with the reported human GINSstructures [53]. Each subunit of human GINS shares a similar fold, and assembles into theheterotetramer of a unique trapezoidal shape [54-56]. Sld5 and Psf1 possess the α-helical (A)domain at the N-terminus and the β-stranded domain (B) at the C-terminus (AB-type). Onthe other hand, Psf2 and Psf3 are the permuted version (BA-type). The backbone structure ofeach subunit and the tetrameric assembly of T. kodakarensis GINS are similar to those of hu‐man GINS. However, the location of the C-terminal B domain of Gins51 is remarkably dif‐ferent between the two GINS structures [53]. A homology model of the homotetramericGINS from T. acidophilum was performed using the T. kodakarensis GINS crystal structure asa template. The Gins 51 protein has a long disordered region inserted between the A and Bdomains and this allows the conformation of the C-terminal domains to be more flexible.


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This domain arrangement leads to the formation of an asymmetric homotetramer, ratherthan a symmetrical assembly, of the T. kodakarensis GINS [53].

The Cdc45 protein is ubiquitously distributed from yeast to human, supporting the notionthat the formation of the CMG complex is universal in the eukaryotic DNA replication proc‐ess. However, no archaeal homologue of Cdc45 has been identified. A recent report of bioin‐formatic analysis showed that the primary structure of eukaryotic Cdc45 and prokaryoticRecJ share a common ancestry [57]. Indeed, a homolog of the DNA binding domain of RecJhas been co-purified with GINS from S. solfataricus [50]. Our experiment detected the stimu‐lation of the 5’-3’ exonuclease activity of the RecJ homologs from P. furiosus and T. kodakaren‐sis by the cognate GINS complexes (Ishino et al., unpublished). The RecJ homolog from T.kodakarensis forms a stable complex with the GINS, and the 5’-3’ exonuclease activity is en‐hanced in vitro; therefore, the RecJ homolog was designated as GAN, from GINS-AssociatedNuclease in a very recent paper [58]. Another related report found that the human Cdc45structure obtained by the small angle X-ray scattering analysis (SAXS) is consistent with thecrystallographic structure of the RecJ family members [59]. These current findings will pro‐mote further research on the structures and functions of the higher-order unwindosome inarchaeal and eukaryotic cells (Figure 3).

7. Primase

To initiate DNA strand synthesis, a primase is required for the synthesis of a short oligonu‐cleotide, as a primer. The DnaG and p48-p58 proteins are the primases in Bacteria and Eu‐karyota, respectively. The p48-p58 primase is further complexed with p180 and p70, to formDNA polymerase α-primase complex. The catalytic subunits of the eukaryotic (p48) andarchaeal primases, share a little, but distinct sequence homology with those of the family XDNA polymerases [60]. The first archaeal primase was identified from Methanococcus janna‐schii, as an ORF with a sequence similar to that of the eukaryotic p48. The gene product ex‐hibited DNA polymerase activity and was able to synthesize oligonucleotides on thetemplate DNA [61]. We characterized the p48-like protein (p41) from P. furiosus. Unexpect‐edly, the archaeal p41 protein did not synthesize short RNA by itself, but preferentially uti‐lized deoxynucleotides to synthesize DNA strands up to several kilobases in length [62].Furthermore, the gene neighboring the p41 gene encodes a protein with very weak similari‐ty to the p58 subunit of the eukaryotic primase. The gene product, designated p46, actuallyforms a stable complex with p41, and the complex can synthesize a short RNA primer, aswell as DNA strands of several hundred nucleotides in vitro [63]. The short RNA but notDNA primers were identified in Pyrococcus cells, and therefore, some mechanism to domi‐nantly use RNA primers exists in the cells [10].

Further research on the primase homologs from S. solfataricus [64-66], Pyrococcus horikoshii[67-69], and P. abyssi [70] showed similar properties in vitro. Notably, p41 is the catalytic sub‐unit, and the large one modulates the activity in the heterodimeric archaeal primases. Thesmall and large subunits are also called PriS and PriL, respectively. The crystal structure of


the N-terminal domain of PriL complexed with PriS of S. solfataricus primase revealed thatPriL does not directly contact the active site of PriS, and therefore, the large subunit may in‐teract with the synthesized primer, to adjust its length to a 7-14 mer. The structure of thecatalytic center is similar to those of the family X DNA polymerases. The 3’-terminal nucleo‐tidyl transferase activity, detected in the S. solfataricus primase [64, 66], and the gap-fillingand strand-displacement activities in the P. abyssi primase [70] also support the structuralsimilarity between PriS and the family X DNA polymerases.

A unique activity, named PADT (template-dependent Polymerization Across DiscontinuousTemplate), in the S. solfataricus PriSL complex was published very recently [71]. The activitymay be involved in double-strand break repair in Archaea.

The archaeal genomes also encode a sequence similar to the bacterial type DnaG primase.The DnaG homolog from the P. furiosus genome was expressed in E. coli, but the protein didnot show any primer synthesis activity in vitro, and thus the archaeal DnaG-like protein maynot act as a primase in Pyrococcus cells (Fujikane et al. unpublished). The DnaG-like proteinwas shown to participate in RNA degradation, as an exosome component [72, 73]. However,a recent paper reported that a DnaG homolog from S. solfataricus actually synthesizes pri‐mers with a 13 nucleotide length [74]. It would be interesting to investigate if the two differ‐ent primases share the primer synthesis for leading and lagging strand replication,respectively, in the Sulfolobus cells, as the authors suggested [74]. A proposed hypothesisabout the evolution of PriSL and DnaG from the last universal common ancestor (LUCA) isinteresting [71].

The Sulfolobus PriSL protein was shown to interact with Mcm through Gins23 [50]. This pri‐mase-helicase interaction probably ensures the coupling of DNA unwinding and primingduring the replication fork progression [50]. Furthermore, the direct interaction betweenPriSL and the clamp loader RFC (described below) in S. solfataricus may regulate the primersynthesis and its transfer to DNA polymerase in archaeal cells [75].

8. Single-stranded DNA binding protein

The single-stranded DNA binding protein, which is called SSB in Bacteria and RPA in Arch‐aea and Eukaryota, is an important factor to protect the unwound single-stranded DNAfrom nuclease attack, chemical modification, and other disruptions during the DNA replica‐tion and repair processes. SSB and RPA have a structurally similar domain containing acommon fold, called the OB (oligonucleotide/oligosaccharide binding)-fold, although thereis little amino acid sequence similarity between them [76]. The common structure suggeststhat the mechanism of single-stranded DNA binding is conserved in living organisms de‐spite the lack of sequence similarity. E. coli SSB is a homotetramer of a 20 kDa peptide withone OB-fold, and the SSBs from Deinococcus radiodurans and Thermus aquaticus consist of ahomodimer of the peptide containing two OB-folds. The eukaryotic RPA is a stable hetero‐trimer, composed of 70, 32, and 14 kDa proteins. RPA70 contains two tandem repeats of anOB-fold, which are responsible for the major interaction with a single-stranded DNA in its


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central region. The N-terminal and C-terminal regions of RPA70 mediate interactions withRPA32 and also with many cellular or viral proteins [77, 78]. RPA32 contains an OB-fold inthe central region [79-81], and the C-terminal region interacts with other RPA subunits andvarious cellular proteins [77, 78. 82, 83]. RPA14 also contains an OB-fold [77]. The eukaryoticRPA interacts with the SV40 T-antigen and the DNA polymerase α-primase complex, andthus forms part of the initiation complex at the replication origin [84]. The RPA also stimu‐lates Polα-primase activity and PCNA-dependent Pol δ activity [85, 86].

The RPAs from M. jannaschii and M. thermautotrophicus were reported in 1998, as the first arch‐aeal single-stranded DNA binding proteins [87-89]. These proteins share amino acid sequencesimilarity with the eukaryotic RPA70, and contain four or five repeated OB-fold and one zinc-finger motif. The M. jannaschii RPA exists as a monomer in solution, and has single-strandDNA binding activity. On the other hand, P. furiosus RPA forms a complex consisting of threedistinct subunits, RPA41, RPA32, and RPA14, similar to the eukaryotic RPA [90]. The P. furio‐sus RPA strikingly stimulates the RadA-promoted strand-exchange reaction in vitro [90].

While the euryarchaeal organisms have a eukaryotic-type RPA homologue, the crenarchaealSSB proteins appear to be much more related to the bacterial proteins, with a single OB foldand a flexible C-terminal tail. However, the crystal structure of the SSB protein from S. solfa‐taricus showed that the OB-fold domain is more similar to that of the eukaryotic RPAs, sup‐porting the close relationship between Archaea and Eukaryota [91].

The RPA from Methanosarcina acetivorans displays a unique property. Unlike the multipleRPA proteins found in other archaea and eukaryotes, each subunit of the M. acetivoransRPAs, RPA1, RPA2, and RPA3, have 4, 2, and 2 OB-folds, respectively, and can act as a dis‐tinct single-stranded DNA-binding proteins. Furthermore, each of the three RPA proteins,as well as their combinations, clearly stimulates the primer extension activity of M. acetivor‐ans DNA polymerase BI in vitro, as shown previously for bacterial SSB and eukaryotic RPA[92]. Architectures of SSB and RPA suggested that they are composed of different combina‐tions of the OB fold. Bacterial and eukaryotic organisms contain one type of SSB or RPA, re‐spectively. In contrast, archaeal organisms have various RPAs, composed of differentorganizations of OB-folds. A hypothesis that homologous recombination might play an im‐portant role in generating this diversity of OB-folds in archaeal cells was proposed, based onexperiments characterizing the engineered RPAs with various OB-folds [93].

9. DNA polymerase

DNA polymerase catalyzes phosphodiester bond formation between the terminal 3’-OH ofthe primer and the α-phosphate of the incoming triphosphate to extend the short primer,and is therefore the main player of the DNA replication process. Based on the amino acidsequence similarity, DNA polymerases have been classified into seven families, A, B, C, D,E, X, and Y (Table 2) [94-98].

The fundamental ability of DNA polymerases to synthesize a deoxyribonucleotide chain iswidely conserved, but more specific properties, including processivity, synthesis accuracy,


and substrate nucleotide selectivity, differ depending on the family. The enzymes within thesame family have basically similar properties. E. coli has five DNA polymerases, and Pol I,Pol II, and Pol III belong to families A, B, and C, respectively. Pol IV and Pol V are classifiedin family Y, as the DNA polymerases for translesion synthesis (TLS). In eukaryotes, the rep‐licative DNA polymerases, Pol α, Pol δ, and Pol ε, belong to family B, and the translesionDNA polymerases, η, ι, and κ, belong to family Y [99].

The most interesting feature discovered at the inception of this research area was that thearchaea indeed have the eukaryotic Pol α-like (Family B) DNA polymerases [100-102]. Mem‐bers of the Crenarchaeota have at least two family B DNA polymerases [103, 104]. On theother hand, there is only one family B DNA polymerase in the Euryarchaeota. Instead, theeuryarchaeal genomes encode a family D DNA polymerase, proposed as Pol D, whichseems to be specific for these archaeal organisms and has never been found in other do‐mains [95, 105]. The genes for family Y-like DNA polymerases are conserved in several, butnot all, archaeal genomes. The role of each DNA polymerase in the archaeal cells is still notknown, although the distribution of the DNA polymerases is getting clearer (Table 2) [106].

5

families of DNA polymerases

A B C D E X Y

Archaea

Crenarchaeota Pol BI, Pol BII Pol BIII

Pol E* Pol Y

Euryarchaeota Pol BI Pol D Pol E* Pol Y

Korarchaeota Pol BI, Pol BII Pol D

Aigarchaeota Pol BI, Pol BII Pol D Pol Y

Thaumarchaeota Pol BI Pol D Pol Y

Bacteria Pol I Pol II Pol III Pol IV Pol V

Eukaryota Pol θ Pol γ**

Pol α, Pol δ Pol ε, Pol ζ

Pol β, Pol λ Pol μ, Pol σ

Pol η, Pol ι Pol κ

* plasmid-encoded

** mitochondrial

Table 2. Distribution of DNA polymerases from seven families in the three domains of life.


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The first family D DNA polymerase was identified from P. furiosus, by screening for DNApolymerase activity in the cell extract [107]. The corresponding gene was cloned, revealingthat this new DNA polymerase consists of two proteins, named DP1 and DP2, and that thededuced amino acid sequences of these proteins were not conserved in the DNA polymer‐ase families [8]. P. furiosus Pol D exhibits efficient strand extension activity and strong proof-reading activity [8, 108]. Other family D DNA polymerases were also characterized byseveral groups [109-115]. The Pol D genes had been found only in Euryarchaeota. However,recent environmental genomics and cultivation efforts revealed novel phyla in Archaea:Thaumarchaeota, Korarchaeota, and Aigarchaeota, and their genome sequences harbor thegenes encoding Pol D.

A genetic study on Halobacterium sp. NRC-1 showed that both Pol B and Pol D are essentialfor viability [116]. An interesting issue is to elucidate whether Pol B and Pol D work togetherat the replication fork for the synthesis of the leading and lagging strands, respectively. Ac‐cording to the usage of an RNA primer and the presence of strand displacement activity, PolD may catalyze lagging strand synthesis [106, 114].

Thaumarchaeota and Aigarchaeota harbor the genes encoding Pol D and crenarchaeal PolBII [117, 118], while Korarchaeota encodes Pol BI, Pol BII and Pol D [119]. Biochemical char‐acterization of these gene products will contribute to research on the evolution of DNA pol‐ymerases in living organisms. A hypothesis that the archaeal ancestor of eukaryotesencoded three DNA polymerases, two distinct family B DNA polymerases and a family DDNA polymerase, which all contributed to the evolution of the eukaryotic replication ma‐chinery, consisting of Pol α, δ, and ε, has been proposed [120].

A protein is encoded in the plasmid pRN1 isolated from a Sulfolobus strain [121]. This pro‐tein, ORF904 (named RepA), has primase and DNA polymerase activities in the N-terminaldomain and helicase activity in the C-terminal domain, and is likely to be essential for thereplication of pRN1 [122, 123]. The amino acid sequence of the N-terminal domain lacks ho‐mology to any known DNA polymerases or primases, and therefore, family E is proposed.Similar proteins are encoded by various archaeal and bacterial plasmids, as well as by somebacterial viruses [124]. Recently, one protein, tn2-12p, encoded in the plasmid pTN2 isolatedfrom Thermococcus nautilus, was experimentally identified as a DNA polymerase in this fam‐ily [125]. This enzyme is likely responsible for the replication of the plasmids. Further inves‐tigations of this family of DNA polymerases will be interesting from an evolutionalperspective.

10. PCNA and RFC

The sliding clamp with the doughnut-shaped ring structure is conserved among living or‐ganisms, and functions as a platform or scaffold for proteins to work on the DNA strands.The eukaryotic and archaeal PCNAs form a homotrimeric ring structure [126, 127], whichencircles the DNA strand and anchors many important proteins involved in DNA replica‐tion and repair (Figure 4). PCNA works as a processivity factor that retains the DNA poly‐


merase on the DNA by binding it on one surface (front side) of the ring for continuous DNAstrand synthesis in DNA replication (Figure 5). To introduce the DNA strand into the centralhole of the clamp ring, a clamp loader is required to interact with the clamp and open itsring. The archaeal and eukaryotic clamp loader is called RFC (Figure 5). The most studiedarchaeal PCNA and RFC molecules to date are P. furiosus PCNA [128-132] and RFC[133-136]. The PCNA and RFC molecules are essential for DNA polymerase to performprocessive DNA synthesis. The molecular mechanism of the clamp loading process has beenactively investigated [137] (Figure 5). An intermediate PCNA-RFC-DNA complex, in whichthe PCNA ring is opened with out-of plane mode, was detected by a single particle analysisof electron microscopic images using P. furiosus proteins (Figure 6) [138]. The crystal struc‐ture of the complex, including the ATP-bound clamp loader, the ring-opened clamp, and thetemplate-primer DNA, using proteins from bacteriophage T4, has recently been published[139], and our knowledge about the clamp loading mechanism is continuously progressing.

Figure 4. PCNA-interacting proteins

After clamp loading, DNA polymerase accesses the clamp and the polymerase-clamp com‐plex performs processive DNA synthesis. Therefore, structural and functional analyses ofthe DNA polymerase-PCNA complex is the next target to elucidate the overall mechanismsof replication fork progression. The PCNA interacting proteins contain a small conserved se‐quence motif, called the PIP box, which binds to a common site on PCNA [140]. The PIP boxconsists of the sequence “Qxxhxxaa”, where “x” represents any amino acid, “h” represents ahydrophobic residue (e.g. L, I or M), and “a” represents an aromatic residue (e.g. F, Y or W).Archaeal DNA polymerases have PIP box-like motifs in their sequences [141]). However,only a few studies have experimentally investigated the function of the motifs. The crystalstructure of P. furiosus Pol B complexed with a monomeric PCNA mutant was determined,


105

and a convincing model of the polymerase-PCNA ring interaction was constructed [142].This study revealed that a novel interaction is formed between a stretched loop of PCNAand the thumb domain of Pol B, in addition to the authentic PIP box. A comparison of themodel structure with the previously reported structures of a family B DNA polymerasefrom RB69 phage, complexed with DNA [143, 144], suggested that the second interactionsite plays a crucial role in switching between the polymerase and exonuclease modes, by in‐ducing a PCNA-polymerase complex configuration that favors synthesis over editing. Thisputative mechanism for the fidelity control of replicative DNA polymerases is supported byexperiments, in which mutations at the second interaction site enhanced the exonuclease ac‐tivity in the presence of PCNA [144]. Furthermore, the three-dimensional structure of theDNA polymerase-PCNA-DNA ternary complex was analyzed by electron microscopic (EM)single particle analysis. This structural view revealed the entire domain configuration of thetrimeric ring of PCNA and DNA polymerase, including the protein-protein or protein-DNAcontacts. This architecture provides clearer insights into the switching mechanism betweenthe editing and synthesis modes [145].

Figure 5. Mechanisms of processive DNA synthesis The clamp loader (RFC) tethers the clamp (PCNA) onto the pri‐mer terminus of the DNA strand. The clamp loader is then replaced by DNA polymerase, which can synthesize theDNA strand processively without falling off.

In contrast to most euryarchaeal organisms, which have a single PCNA homolog forming ahomotrimeric ring structure, the majority of crenarchaea have multiple PCNA homologues,and they are capable of forming heterotrimeric rings for their functions [146, 147]. It is espe‐cially interesting that the three PCNAs, PCNA1, PCNA2, and PCNA3, specifically bindPCNA binding proteins, including DNA polymerases, DNA ligases, and FEN-1 endonu‐clease [147, 148]. Detailed structural studies of the heterologous PCNA from S. solfataricusrevealed that the interaction modes between the subunits are conserved with those of thehomotrimeric PCNAs [149, 150].

T. kodakarensis is the only euryarchaeal species that has two genes encoding PCNA homo‐logs on the genome [38]. These two genes from the T. kodakarensis genome, and the highlypurified gene products, PCNA1 and PCNA2, were characterized [151]. PCNA1 stimulatedthe DNA synthesis reactions of the two DNA polymerases, Pol B and Pol D, from T. kodakar‐ensis in vitro. PCNA2 however only had an effect on Pol B. The T. kodakarensis strain withpcna2 disruption was isolated, whereas gene disruption for pcna1 was not possible. These re‐sults suggested that PCNA1 is essential for DNA replication, and PCNA2 may play a differ‐ent role in T. kodakarensis cells. The sensitivities of the Δpcna2 mutant strain to ultraviolet


irradiation (UV), methyl methanesulfonate (MMS) and mitomycin C (MMC) were indistin‐guishable to those of the wild type strain. Both PCNA1 and PCNA2 form a stable ring struc‐ture and work as a processivity factor for T. kodakarensis Pol B in vitro. The crystal structuresof the two PCNAs revealed the different interactions at the subunit-subunit interfaces [152].

Figure 6. Electron Microscopic Analysis of P. furious DNA polymerase-PCNA-DNA complex. The complex in theediting mode of the DNA polymerase-PCNA-DNA ternary complex was shown. (A) Electron microscopic (EM) map ofthe complex is depicted by gray surface. DNA polymerase and PCNA are shown in a ribbon representation coloredpurple and blue, respectively. The DNA is colored white. The exonuclease active site is shown in a green ribbon. (B)Schematic view of the complex.

The RFC molecule is conserved as a pentameric complex in Eukaryota and Archaea. Howev‐er, the eukaryotic RFC is a heteropentameric complex, consisting of five different proteins,RFC1 to 5, in which RFC1 is larger than the other four RFCs. On the other hand, the archaealRFC consists of two proteins, RFCS (small) and RFCL (large), in a 4 to 1 ratio. A differentform of RFC, consisting of three subunits, RFCS1, RFCS2, and RFCL, in a 3 to 1 to 1 ratio,was also identified from M. acetivorans [153]. The three subunits of RFC may represent anintermediate stage in the evolution of the more complex RFC in Eukaryota from the lesscomplex RFC in Archaea [153, 154]. The subunit organization and the spatial distribution ofthe subunits in the M. acetivorans RFC complex were analyzed and compared with those ofthe E. coli γ-complex, which is also a pentamer consisting of three different proteins. Thesetwo clamp loaders adopt similar subunit organizations and spatial distributions, but thefunctions of the individual subunits are likely to be diverse [154].

11. DNA ligase

DNA ligase is essential to connect the Okazaki fragments of the discontinuous strand syn‐thesis during DNA replication, and therefore, it universally exists in all living organisms.This enzyme catalyzes phosphodiester bond formation via three nucleotidyl transfer steps[155, 156]. In the first step, DNA ligase forms a covalent enzyme-AMP intermediate, by re‐acting with ATP or NAD+ as a cofactor. In the second step, DNA ligase recognizes the sub‐strate DNA, and the AMP is subsequently transferred from the ligase to the 5’-phosphateterminus of the DNA, to form a DNA-adenylate intermediate (AppDNA). In the final step,


107

the 5’-AppDNA is attacked by the adjacent 3’-hydroxy group of the DNA and a phospho‐diester bond is formed. DNA ligases are grouped into two families, according to their re‐quirement for ATP or NAD+ as a nucleotide cofactor in the first step reaction. ATP-dependent DNA ligases are widely found in all three domains of life, whereas NAD+-dependent DNA ligases exist mostly in Bacteria. Some halophilic archaea [157] andeukaryotic viruses [158] also have NAD+-dependent enzymes.

Three genes (LIG1, LIG3 and LIG4) encoding ATP-dependent DNA ligases have been identi‐fied in the human genome to date and DNA ligase I (Lig I), encoded by LIG1, is a replicativeenzyme that joins Okazaki fragments during DNA replication [156]. The first gene encodinga eukaryotic-like ATP-dependent DNA ligase was found in the thermophilic archaeon, De‐sulfolobus ambivalens [159]. Subsequent identifications of the DNA ligases from archaeal or‐ganisms revealed that these enzymes primarily use ATP as a cofactor. However, thisclassification may not be so strict. The utilization of NAD+, as well as ATP, as a cofactor hasbeen observed in several DNA ligases, including those from T. kodakarensis [160], T. fumico‐lans, P. abyssi [161]), Thermococcus sp. NA1 [162], T. acidophilum, Picrophilus torridus, and Fer‐roplasma acidophilum, although ATP is evidently preferable in all of the cases [163] (Table 3).The dual co-factor specificity (ATP/NAD+) is an interesting feature of these DNA ligase en‐zymes and it will be enlightening to investigate the structural basis for this. Another dualco-factor specificity exists in the archaeal DNA ligases, which use ADP as well as ATP, asfound in the enzymes from A. pernix [164] and Staphylothermus marinus [165], and in the caseof Sulfobococcus zilligii, GTP is also the functional cofactor [166]. The DNA ligases from P.horikoshii [167] and P. furiosus [168] have a strict ATP preference (Table 3). Sufficient bio‐chemical data have not been obtained to resolve the issue of dual co-factor specificity, andfurther biochemical and structural analyses are required.

cofactor

ATP ATP and ADP ATP and NAD+ ATP, ADP, and GTP

Acidithiobacillus ferrooxidans Aeropyrum pernix Ferroplasma acidophilum Sulfophobococcus zilligii

Ferroplasma acidarmanus Staphylothermus marinus Picrophilus torridus

Methanothermobacterium

thermoautotrophicum

Thermococcus fumicolans

Pyrococcus horikoshii Thermococcus kodakarensis

Pyrococcus furiosus Thermococcus sp.

Sulfolobus acidocaldarius Thermoplasma acidophilum

Sulfolobus shibatae

Thermococcus sp. 1519

Table 3. Cofactor dependency of the archaeal DNA ligases


The crystal structure of P. furiosus DNA ligase [169] was solved and the physical and func‐tional interactions between the DNA ligase and PCNA was shown [168]. The detailed inter‐action mode between human Lig I and PCNA is somewhat unclear, because of severalcontroversial reports [170-172]. The stimulatory effect of P. furiosus PCNA on the enzyme ac‐tivity of the cognate DNA ligase was observed at a high salt concentration, at which a DNAligase alone cannot bind to a nicked DNA substrate. Interestingly, the PCNA-binding site islocated in the middle of the N-terminal DNA binding domain (DBD) of the P. furiosus DNAligase, and the binding motif, QKSFF, which is proposed as a shorter version of the PIP box,is actually looped out from the protein surface [168]. Interestingly, this motif is located in themiddle of the protein chain, rather than the N- or C-terminal region, where the PIP boxes areusually located. To confirm that this motif is conserved in the archaeal/eukaryotic DNA li‐gases, the physical and functional interactions between A. pernix DNA ligase and PCNAwas analyzed and the interaction was shown to mainly depend on the phenylalanine 132residue, which is located in the predicted region from the multiple sequence alignment ofthe ATP-dependent DNA ligases [173].

The crystal structure of the human Lig I, complexed with DNA, was solved as the first ATP-dependent mammalian DNA ligase, although the ligase was an N-terminal truncated form[174]. The structure comprises the N-terminal DNA binding domain, the middle adenyla‐tion domain, and the C-terminal OB-fold domain. The crystal structure of Lig I (residues 233to 919) in complex with a nicked, 5'-adenylated DNA intermediate revealed that the enzymeredirects the path of the dsDNA, to expose the nick termini for the strand-joining reaction.The N-terminal DNA-binding domain works to encircle the DNA substrate like PCNA andto stabilize the DNA in a distorted structure, positioning the catalytic core on the nick. Thecrystal structure of the full length DNA ligase from P. furiosus revealed that the architectureof each domain resembles those of Lig I, but the domain arrangements strikingly differ be‐tween the two enzymes [168]. This domain rearrangement is probably derived from the “do‐main-connecting” role of the helical extension conserved at the C-termini in the archaeal andeukaryotic DNA ligases. The DNA substrate in the open form of Lig I is replaced by motifVI at the C-terminus, in the closed form of P. furiosus DNA ligase. Both the shapes and elec‐trostatic distributions are similar between motif VI and the DNA substrate, suggesting thatmotif VI in the closed state mimics the incoming substrate DNA. The subsequently solvedcrystal structure of S. solfataricus DNA ligase is the fully open structure, in which the threedomains are highly extended [175]. In this work, the S. solfataricus ligase-PCNA complexwas also analyzed by SAXS. S. solfataricus DNA ligase bound to the PCNA ring still retainsan open, extended conformation. The closed, ring-shaped conformation observed in the LigI structure as described above is probably the active form to catalyze a DNA end-joining re‐action, and therefore, it is proposed that the open-to-closed movement occurs for ligation,and the switch in the conformational change is accommodated by a malleable interface withPCNA, which serves as an efficient platform for DNA ligation [175]. After the publication ofthese crystal structures, the three-dimensional structure of the ternary complex, consistingof DNA ligase-PCNA-DNA, using the P. furiosus proteins was obtained by EM single parti‐cle analysis [176]. In the complex structure, the three domains of the crescent-shaped P. fur‐iosus DNA ligase surround the central DNA duplex, encircled by the closed PCNA ring. The


109

relative orientations of the ligase domains remarkably differ from those of the crystal struc‐tures, and therefore, a large domain rearrangement occurs upon ternary complex formation.In the EM image model, the DNA ligase contacts PCNA at two sites, the conventional PIPbox and a novel second contact in the middle adenylation domain. It is also interesting thata substantial DNA tilt from the PCNA ring axis is observed. Based on these structural analy‐ses, a mechanism in which the PCNA binding proteins are bound and released sequentially.In fact, most of the PCNA binding proteins share the same binding sites in the interdomainconnecting loop (IDCL) and the C-terminal tail of the PCNA. The structural features excludethe possibility that the three proteins contact the single PCNA ring simultaneously, becauseDNA ligase occupies two of the three subunits of the PCNA trimer. In the case of the RFC-PCNA-DNA complex structure obtained by the same EM technique, RFC entirely covers thePCNA ring, thus blocking the access of other proteins [138]. These ternary complexes appearto favor a mechanism involving the sequential binding and release of replication factors.

12. Flap endonuclease 1 (FEN1)

Efficient processing of Okazaki fragments to make a continuous DNA strand is essential forthe lagging strand synthesis in asymmetric DNA replication. The primase-synthesizedRNA/DNA primers need to be removed to join the Okazaki fragments into an intact contin‐uous strand DNA. Flap endonuclease 1 (FEN1) is mainly responsible for this task. Okazakifragment maturation is highly coordinated with continuous DNA synthesis, and the interac‐tions of DNA polymerase, FEN1, and DNA ligase with PCNA allow these enzymes to actsequentially during the maturation process, as described above.

FEN1, a structure-specific 5’-endonuclease, specifically recognizes a dsDNA with an unan‐nealed 5’-flap [177, 178]. In the eukaryotic Okazaki fragment processing system, 5’-flapDNA structures are formed by the strand displacement activity of DNA polymerase δ. Lig Iseals the nick after the flapped DNA is cleaved by FEN1. These processing steps are facilitat‐ed by PCNA [179]. The interactions between eukaryotic FEN1 and PCNA have been wellcharacterized [140, 171], and the stimulatory effect of PCNA on the FEN1 activity was alsoshown [180]. The crystal structure of the human FEN1-PCNA complex revealed three FEN1molecules bound to each PCNA subunit of the trimer ring in different configurations [181].Based on these structural analyses together with the description in the DNA ligase section, aflip-flop transition mechanism, which enables proteins to internally switch for differentfunctions on the same DNA clamp are currently being considered.

The eukaryotic homologs of FEN1 were found in Archaea [182]. The crystal structures ofFEN1 from M. jannaschii [183], P. furiosus [184], P. horikoshii [185], A. fulgidus [186], and S.solfataricus [150] have been determined. In addition, detailed biochemical studies were per‐formed on P. horikoshii FEN1 [187, 188]. Thus, studies of the archaeal FEN1 proteins haveprovided important insights into the structural basis of the cleavage reaction of the flappedDNA. Our recent research showed that the flap endonuclease activity of P. furiosus FEN1was stimulated by PCNA. Furthermore, the stimulatory effect of PCNA on the sequential


action of FEN1 and DNA ligase was observed in vitro (Kiyonari et al., unpublished). Basedon these results, a model of the molecular switching mechanisms of the last steps of Okaza‐ki-fragment maturation was constructed. The quaternary complex of FEN1-Lig-PCNA-DNAwas also isolated for the EM single particle analysis. These studies will provide more con‐crete image of the molecular mechanism.

13. Summary and perspectives

Research on the molecular mechanism of DNA replication has been a central theme of mo‐lecular biology. Archaeal organisms became popular in the total genome sequencing age, asdescribed above, and most of the DNA replication proteins are now equally understood bybiochemical characterizations. In addition, the archaeal studies are especially interesting tounderstand the mechanisms by which cells live in extreme environmental conditions. Fur‐thermore, it is also noteworthy that the proteins from the hyperthermophilic archaea aremore stable than those from mesophilic organisms, and they are advantageous for the struc‐tural and functional analyses of higher-ordered complexes, such as the replisome. Studieson the higher-ordered complexes, rather than single proteins, are essential for understand‐ing each of the events involved in DNA metabolism, and the archaeal research will continu‐ously contribute to the development and advancement of the DNA replication researchfield, as summarized in part in a recent review [189, 190].

In addition to basic molecular biology research, DNA replication proteins from thermo‐philes have been quite useful reagents for gene manipulations, including genetic diagnosis,forensic DNA typing, and detection of bacterial and virus infections, as well as basic re‐search. Numerous enzymes have been commercialized around the world, and are utilizeddaily. An example of the successful engineering of an archaeal DNA polymerase for PCR isthe creation of the fusion protein between P. furiosus Pol B and a nonspecific dsDNA bind‐ing protein, Sso7d, from S. solfataricus, by genetic engineering techniques [191]. The fusionDNA polymerase overcame the low processivity of the wild type Pol B by the high affinitySso7d to the DNA strand. As another example, we successfully developed a novel proces‐sive PCR method, using the archaeal Pol B with the help of a mutant PCNA [192, 193]. Sev‐eral DNA sequencing technologies, referred to as “next-generation sequencing”, have beendeveloped [194, 195], and are now commercially available. Single-molecule detection, usingdye-labeled modified nucleotides and longer read lengths, is now known as “third-genera‐tion DNA sequencing” [196]. These technologies apply DNA polymerases or DNA ligasesfrom various sources, indicating that these DNA replication enzymes are indispensable forthe development of DNA manipulation technology. These facts prove that the progress ofthe basic research on the molecular biology of archaeal DNA replication will promote thedevelopment of the new technologies for genetic engineering.


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Author details

Yoshizumi Ishino and Sonoko Ishino

Department of Bioscience and Biotechnology, Graduate School of Bioresource and Bioenvir‐onmental Sciences, Kyushu University, Japan

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